Steinernema sp. nematode for suppression of Helicoverpa zea and Spodoptera frugiperda

ABSTRACT

A novel entomopathogenic nematode of the genus Steinernema, which is effective as a biopesticide for the control of insects, and particularly the corn earworm, Helicoverpa zea, and the fall armyworm, Spodoptera frugiperda. This nematode has been identified as Steinernema riobravis.

BACKGROUND OF THE INVENTION

1. Field of the Invention

The corn earworm (CEW), Helicoverpa zea (also known as Heliothis zea),is a prominant pest of cotton, sorghum, tobacco and numerous vegetablecrops, particularly corn, throughout the world.

This invention relates to a novel entomopathogenic nematode of the genusSteinernema, which is effective as a biopesticide for the control ofinsects, and particularly the corn earworm.

2. Description of the Prior Art

Recent studies have shown that the Lower Rio Grande Valley corn crop isa major source for the production of large populations of the cornearworm and fall armyworm which migrate north to infest corn and othercrops of higher cash value such as cotton, tomatoes and soybeans, wherethey cause severe economic damage [Raulston et al., Production ofHeliothis zea on Corn in Northeastern Mexico and the Lower Rio GrandeValley of Texas: a Potential Source for Corn and Cotton Infestation onthe High Plains of Texas, Proc. Beltwide Cotton Conf., 1986, pp.222-225Memphis, National Cotton Council of America; Wolf et al, Phil.Trans. R. Soc. Lond. B 328:619-630, (1990); and Pair et al, FloridaEntomologist, 74:200-213, (1991)]. The primary control strategy for thecorn earworm is the application of insecticides that result in egg andlarval mortality. In Florida and other southeastern states, insecticidesare usually applied at least every 48 hours to protect sweet corn duringthe silking period. However, the corn earworm has given indications ofresistance to organochlorine and organophosphate insecticides[Wolfenbarger et al., Bull. Entomol. Soc. Amer., 27:181-185, (1981); andSparks, Bull. Entomol. Soc. Amer., 27:186-192, (1981)]. This resistance,in addition to the wide public awareness of the environmental damageresulting from chemical pesticides, has increased interest in biologicalcontrol, and has resulted in the study of a variety of biologicalcontrol agents. Entomopathogenic nematodes have shown promise asbiological control candidates for a number of insect pests.

Nematodes of the genera Steinernema and Heterorhabditis possess most ofthe characteristics of an ideal biological control agent for insects[Poiner, Taxonomy and Biology of Steinernematidae and Heterorhabditidae,In Gaugler and Kaya (eds.), Entomopathogenic Nematodes in BiologicalControl, CRC Press, Boca Raton, Fla., (1990), pages 23-61; and Gaugler,J. Nematol., 13:241-249, (1981)]]. These nematodes search for theirinsect hosts; they are highly virulent, killing most hosts within 48hours; they are easily and inexpensively mass produced; and they have awide range of insect hosts (Poiner, ibid and Gaugler, ibid). Theeffectiveness of these nematodes is attributed to a mutualisticbacterium of the genus Xenorhabdus associated therewith [Poinar, ibid].After entry or penetration of the nematode into the insect host, thebacteria are released from the nematode and rapidly multiply, killingthe host insect by septicemia. Conversely, the nematodes protect thebacteria from the environment prior to release within a suitable host.The pathogenicity of entompathogenic nematodes to Heliothesis specieshas been demonstrated previously [Tanada and Reiner, J. Inverteb. Path.4:139-154, (1962); Bong and Sikorowski, J. Econ. Ent., 76:590-593(1983); and Howell, J. Inverteb. Path., 33:155-158, (1979)]. However,there exists a negative relationship between larval age andsusceptibility to the nematodes [Glazer and Navon, J. Econ. Entomol.,83:1795-1800, (1990); and Samsook and Sikora, Med. Fac. Landbouww.Gent., 46:685-693, (1981)]. Consequently, the use of these nematodes hasbeen against the feeding stages of various insect pests, while their useagainst prepupal or pupal stages of H. zea has been limited.

SUMMARY OF THE INVENTION

We have now discovered a previously unknown entomopathogenic nematode ofthe genus Steinernema, which is effective as a biopesticide for thecontrol of insects, and particularly the corn earworm, Helicoverpa zea,and the fall armyworm, Spodoptera frugiperda. This nematode has beenidentified as Steinernema riobravis.

The nematode of this invention has been isolated in pure form from pupaeand prepupae of the corn earworm form soil samples of corn fields in theLower Rio Grande Valley of Texas. When applied to the locus of thetarget insects, Steinernema riobravis will provide improved suppressionof the insect population.

In accordance with this discovery, it is an object of this invention tointroduce Steinernema riobravis as a novel biopesticide for the controlof insects. It is also an object of this invention to provide newcompositions and methods for controlling insect populationsincorporating Steinernema riobravis.

A further object of this invention is to provide a biopesticide that iseffective against the non-feeding pupal or prepupal stages of insectsand that may be applied to the soil.

Another object of this invention is to provide a biopesticide thatremains viable at relatively low moisture conditions, is effective forcontrolling insects in clay soil types, and that provides effectiveinsect control at low inocolum levels.

Yet another object of this invention is to provide a biopesticide forthe suppression or elimination of the corn earworm and fall armyworminsects at their source, thereby preventing their movement to othercrops.

Other objects and advantages of this invention will become readilyapparent from the ensuring description.

DETAILED DESCRIPTION

The entomopathogenic nematode of this invention, Steinernema riobravis,is indigenous to the Lower Rio Grande Valley of Texas and northernTamaulipas, Mexico and may be recovered from corn fields within thisgeographical area as will be described in more detail hereinbelow.Indeed, over a five year period, 34% of all fields sampled containedcorn earworm parasitized with this nematode, and 24.2% contained fallarmyworm parasitized with the nematode. Of all corn earworm and fallarmyworm prepuae and pupae collected during this study, 11.6% and 9.3%,respectively, were parasitized with the nematode.

The above-mentioned Steinernema riobravis has been deposited under theBudapest Treaty in the American Type Culture Collection (ATCC), 10801University Blvd., Manassas, Va., 20110-2209, USA, on Apr. 1, 1994, andhas been assigned accession no. ATCC 75727.

Steinernema riobravis n.sp. can be separated from other Steinernema spp.by several characters. Included among the separating traits is thelength of the infective juvenile (average length-622 microns;range=561-701 microns). Further the male posterior segment of S.riobravis n.sp. lacks a projection or spine of any type which separatesit from Steinernema carpocapsae and Steinernema feltiae. The spiculesare generally more curved (a line running parallel with the calomus andlamina forms an angle of 90 to 100 degrees) than those of S.carpocapsae, S. feltia and Steinernema glaseri. The blunt tip of thespicules on S. riobravis n.sp. are distinct from the hooked tip spiculesof S. carpocapsae. Digestion of genomic DNA with restrictionendonucleases generated a unique set of different sized DNA restrictionfragments dependent upon the base of the genome sequence. The sizedistribution of these restriction fragments are different from all knownspecies of nematodes. Morphologically, S. riobravis n.sp. resemblesSteinernema intermedia, however, controlled mating studies indicatedthat Steinernema intermedia n.sp. and S. intermedia do not mate witheach other.

The nematode described herein is effective for controlling a variety ofinsects. Without being limited thereto, pests of particular interestknown to be susceptible to treatment are agronomically importantinsects, especially the corn earworm, H. zea, and the fall armyworm, S.frugiperda. The nematode may be applied for the control of theseagronomically important insects on a number of crops, particularly butnot limited to corn, cotton, tomatoes, and soybeans.

Production of the nematode may be accomplished using in vivo or in vitrotechniques known in the art. As described in the Examples herein,Steinernema riobravis may be initially recovered from soil samples takenfrom corn fields in the Lower Rio Grande Valley of Texas and northernTamaulipas, Mexico. Following isolation from the environment, thenematodes may then be reared in vivo in susceptible host insects such asH. zea prepupae or pupae as illustrated in the Examples. However, inaccordance with the preferred embodiment, the nematodes may also beproduced on a large scale using in vitro rearing techniques, such asdescribed by Friedman et al. [Mass Production in Liquid Culture ofInsect-Killing Nematodes, U.S. Pat. No. 5,023,183, issued Jun. 11, 1991,the contents of which are incorporated by reference herein]. Inaccordance with either technique, the nematodes may be subsequentlyharvested and collected in pure or substantially pure form.

Suitable formulations for commercial insecticidal applications would beprepared from nematodes isolated from the environment, particularly invitro cultivated populations of the nematodes, or pure or substantiallypure nematodes. Because of the moisture required by these nematodes forcontinued viability and infectivity, the nematodes are advantageouslyapplied in combination with a suitable inert carrier or vehicle as knownin the art, which carrier is optionally substantially biologically pure.The term “substantially biologically pure inert carrier” is definedherein as an inert carrier having significantly fewer naturallyoccurring microorganisms relative to the environment. The formulationsdescribed herein are storage stable and, depending upon the carrierused, nematode viability can be maintained up to one year withrefrigeration.

As a practical matter, to facilitate handling and transport of thebiopesticide, and to prevent dessication, the formulations of thenematode and carrier should be enclosed within a closed container suchas a drum, jug, flask or plastic bag as is known in the art.

Of particular interest are formulations employing water as a carrier,with a population of the nematodes suspended therein as described in theExamples. In an alternative embodiment the carrier may be a solid phasematerial or encapsulating agent, upon or within which the nematodes canbe immobilized. Suitable carriers of this type include but are notlimited to hydrogel agents such as alginate gels, wheat-gluten matrices,starch matrices, wheat-bran bait pellets, clay particles, polyacrylamidegels, or synthetic polymers as are known in the art. Preferredalternative carriers and methods for immobilizing nematodes aredescribed, for example, in Nelsen (U.S. Pat. Nos. 4,753,799; 4,701,326and 4,615,883 disclosing alginate gels), Connick and Nickle (U.S. patentapplication Ser. No. 07/560,792, filed Jul. 30, 1990, disclosing wheatgluten), Shasha et al. (U.S. Pat. No. 4,859,377 disclosing starchmatrices), and Capinera and Hibbard [J. Agric. Entomol., 4:337-344,(1987) disclosing wheat-bran bait pellets], the contents of each ofwhich are herein incorporated by reference. Formulations of alginategels containing the nematodes provide the added benefit of enhancedviability after storage, while allowing subsequent conversion to anaqueous liquid by dissolution of the alginate with sodium citrate. Whenthe carrier is other than water, sufficient moisture should be providedto ensure viability and infectivity of the nematodes.

Besides the active agent itself, other additives and adjuncts may beformulated into the compositions of the invention. Examples of theseinclude nutrients, humectants, feeding stimulants (phagostimulants), UVprotectants, inert fillers, and dispersants. Humectant materials includebut are not limited to glycerol, sugars such as sucrose, invertemulsions, and cellulose ethers.

In use, an insecticidally effective amount of the nematode of thisinvention is applied to the locus of, or in the vicinity of, the insectsto be controlled. An “insecticidally effective amount” is defined hereinas that quantity of nematode which will result in a significantmortality rate of a test group of insects compared to an untreatedgroup. The actual effective amount may be readily determined by thepractitioner skilled in the art, and may vary with the species of pest,stage of larval development, the type of vehicle or carrier, the periodof treatment, environmental conditions (especially moisture), and otherrelated factors. Without being limited thereto, in accordance with thepreferred embodiment, the nematodes are applied at a concentrationgreater than or equal to about 2.5×10⁴ infective juveniles per m² ofsoil or field, and especially at a concentration greater than or equalto about 1×10⁵ or 2×10⁵ infective juveniles per m² of soil or field.Surprisingly, these inoculum levels are six times less than the levelsof other entomopathogenic nematodes used to control other soil insectpests [Miller and Bedding, Entomophaga, 27:109-114, (1982). In thealternative, the concentration of nematodes to be applied may also bedetermined relative to the density of the target insects, if known. Thenematodes are preferably applied at a concentration greater than orequal to about 10 infective juveniles per insect, and especially at aconcentration greater than or equal to about 100 infective juveniles perinsect.

Because of the moisture requirements of these nematodes, and since thesoil is their natural habitat and the corn earworm larvae drop to theground to pupate as part of their life-cycle, techniques wherein thenematodes are applied to the soil are desirable. In accordance with aparticularly preferred embodiment for use in areas employing irrigation,the nematodes may be admixed with irrigation water prior to or at thetime of irrigation, effectively distributing the nematodes across thefield. In order to maximize insect kill, the application of thenematodes to the soil should be timed to the exit of prepupae, ensuringthe highest rate of parasitism by the nematodes. Best results have beenachieved when Steinernema riobravis are applied when at least about 10%of the corn earworms have exited the corn ears to pupate, and especiallywhen at least about 50% of the corn earworms have developed into largelarvae. Because Steinernema riobravis actively seek out and thenpenetrate and parasitize the target insect pupae and prepupae, feedingby the insect upon the formulation of the nematode is not required.Further, this nematode has the capability of remaining viable andmaintaining its infectivity in clay soil types for more than two monthsafter application, providing the added advantage of residualinsecticidal activity.

Another technique envisioned for soil application employs encapsulatedor pelletized formulations of the nematode. The capsules or pelletscontaining the nematode may be applied to the soil prior to emergence ofthe crop, or in the case of corn just prior to larval exit from the cornear such as by spreading or spraying. Depending upon the carrierselected, the nematodes may be released from the capsules or pellets asthey degrade in the soil (Connick and Nickle, ibid) or upon ingestion bythe insect. Without being limited thereto, soil application of theseencapsulated or pelletized formulation is ideally conducted prior toemergence of the crop or prior to larval exit from the ear.

While soil application techniques are preferred, the formulations of thenematode may also be applied directly upon the crop such as by spraying.Although the nematode may be applied at any time, when corn is to betreated, the nematode is preferably applied while the corn is in thewhorl stage. Applying the nematode at this time takes advantage ofpockets of moisture retained in whorl folds of the corn because at leastsome of the nematodes would land within the folds where they would beprotected from dessication. Further, the nematodes would still beavailable to attack and/or be ingested by the target insects on theplant.

EXAMPLE 1

Nematode Extraction and Culture. A previously unknown nematode of thegenus Steinernema, subsequently identified as S. riobravis, was isolatedfrom soil samples taken from corn plots after harvest. The corn plotswere located at the U.S. Department of Agriculture South Farm inWeslaco, Tex.

H. zea prepupae were used as trap hosts for this experiment.Approximately 1 kg of a Hidalog sandy loam soil, was collected at eachsample site from the top 10-15 cm of soil. Five prepupae were placed atthe bottom of a 30-cm diam ceramic pot, covered with moist soilexcavated from the corn plots, and incubated at about 23° C. for 7 d.Dead prepupae were transferred to White traps (White, Science,66:302-303, 1929) and infective juveniles (IJ) of the nematode werecollected 10-14 d after exposure to the soil sample.

The Steinernema nematodes were cultured in vivo in the laboratory usingH. zea prepupae as a susceptible host. Following harvest the nematodeswere suspended in 50 ml of water and stored in 275-ml canted neckCorning tissue culture flasks at 10° C. Nematodes were used forexperiments within 1 wk of harvesting.

Helicoverpa Rearing. H. zea, was reared in the laboratory on artificialdiet following the methods of Raulston & Lingren, the contents of whichare incorporated by reference, [Methods for Large-Scale Rearing ofTobacco Budworm, Production Research Report No. 145, ARS, U.S. Dept. ofAgriculture, Washington, D.C., 10 pp, 1972] at 29.5° C. Prepupae used inthis test were 11 days old and an average weight of 644 mg.

EXAMPLE 2

Infectivity Tests. Dosage mortality trials were performed underlaboratory conditions. A 0.5-ml nematode suspension from Example 1 wasadded to filter paper (Whatman No. 1) contained in a petri dish (60×15mm). One prepupa of H. zea was placed in each of four dishes pertreatment, and incubated in the dark, at room temperature (23° C.) for 5d. Dead insects were individually transferred to White trap dishes(White, ibid) to observe nematode development, evaluate insectmortality, and estimate the number of nematodes developed per insect.Nematodes were collected for identification based on morphologicalcharacteristics of the infective juveniles and males (Poinar, ibid).

Dosage morality tests consisted of seven concentrations: 1, 5, 10, 20,40, 80, and 100 nematodes per prepupa. Control prepupae were treatedwith 0.5-ml sterile distilled water alone.

The experiment was designed as a randomized complete block with eighttreatments and four plates per treatment; the experiment was repeatedfive times. The insect mortality data was analyzed by a probit procedureusing the Statistical Analysis System software (SAS Institute, Cary,N.C., 1988). Log₁₀ of the dose value was used in the statisticalanalysis. LD values and 95% Fiducial limits for nematode concentrationswere computed.

Nematode concentrations differentially affected the insect morality ofH. zea prepupae (P=0.0001). One hundred percent mortality of H. zeaprepupae was achieved with exposure to 100 IJ per prepupae (Table 1). Atthis concentration, 60% died in the prepupal stage; however, 40% of theprepupae continued development to the pupal stage prior tonematode-induced death. Doses of 10, 20, 40, and 80 nematodes per H. zeaprepupa caused mortalities of 40, 55, 85, and 90% respectively. Thelowest mortalities (5 and 20%) occurred when prepupae were exposed toonly 1 or 5 nematodes per prepupa. The effective dosage required tocause 50% insect mortality (LD₅₀) was 13 IJ nematodes (Table 1). Thegeneral response of insect mortality (Y) of H. zea as a function ofnematode concentration (N) per prepupa was estimated by a Probitregression model: Probit Y=−2.1+1.8 Log₁₀N (P=0.0001, df=6, andSy.x=0.078).

Production of Nematodes. Fourteen days after nematode infection,nematodes were collected by washing the IJ nematodes contained in the“White trap” dishes and those from the filter paper through a 25-meshscreen sieve into a 2,000 ml plastic beaker. Nematodes were extractedfrom each infected host by transferring a dead insect to a 50-ml plasticcentrifuge tube containing about 5 ml water, grinding it with a spatula,and hand stirring on a magnetic mixer for about 1 min. The tube contentswere washed through the 25-mesh sieve into the beaker. The total volumewas adjusted to 1,000 ml and stirred on a magnetic stir-plate tomaintain a homogeneous suspension of nematodes. A 1-ml aliquot of thesuspension was placed in each of three counting dishes and the averagenumber of nematodes was estimated from counts made under a dissectingmicroscope.

Production of nematodes per prepupa and pupa cadavers was also affectedby the concentration of nematodes to which they were exposed (Table 2).The maximum yield of nematodes/prepupa or pupa occurred at an exposureconcentration of 40 IJ nematodes/prepupa. The least yield of nematodeswas obtained at an exposure level of 5 nematodes/prepupa. The productionof nematodes was not significantly higher on infected prepupae comparedto insects that continued development to the pupal stage. The averagenumbers of nematodes produced per prepupa and pupa cadavers were 325,000and 310,000 respectively.

EXAMPLE 3

Greenhouse Experiment. The greenhouse experiment was designed todetermine the effects of dose and method of Steinernema riobravisapplication onto the soil on the control of H. zea prepupae. Thisexperiment was conducted at the Subtropical Agriculture ResearchLaboratory, U.S. Department of Agriculture, Agriculture Research Servicein Weslaco, Tex. The soil used in this trial was a clay type (51.2%clay, 35.3% sand, and 13.5% silt, 0.5% organic matter, 7.8 pH, 39.95meq/100 g CEC), which was collected from a corn field where thisnematode occurs naturally in the soil. The soil was steam-sterilized,sieved through a 9-mesh sieve, and transferred to 15-cm diam clay pots(1175 cc soil/pot). The greenhouse temperature was about 24±3° and 40%relative humidity. Treatments were arranged in a 5×2 factorial with tentreatment combinations of five doses of nematodes (0, 1250, 2500, 5000,and 10,000 nematodes/pot) and two methods of application (soil surfaceand soil subsurface). The infective juvenile Steinernema riobravis fromExample 1 were stored at 10° C. for two wk and applied to the soil atevening time after 1630 hours CST.

For the first method (soil surface), 10 ml of Steinernema riobravissuspended in sterile distilled water from Example 1 was applied to thesoil surface previously moistened with 350 ml distilled water. Ten H.zea prepupae were evenly distributed on the soil surface in each potafter nematode application. Each H. zea was contained in inverted 20 mlplastic cups with the bottom open to allow the prepupae to burrow in thesoil to pupate. Pots were covered with plastic and small openings (1 mm)were made on the top to reduce evaporation. Five days after nematodetreatment, H. zea were carefully extracted from soil, rinsed andtransferred to “White” trap dishes (white, ibid) to observe nematodedevelopment on dead insects. Evaluation of insect mortality wasperformed 14 d after nematode treatment based on the development ofinfective juvenile nematodes from infected H. zea. Nematodes werecollected for their identification based on morphologicalcharacteristics of infective juveniles and males (Poinar, ibid).

For the second method of application (soil subsurface), the procedurewas similar to the first method except that the Steinernema riobravisnematodes were applied to the soil in two parts. Ten ml of nematodeaqueous suspension containing half of the nematodes was applied to thesoil 2.5 cm deep, then ten H. zea prepupae were placed on the soil ineach pot after nematode treatment and covered with the top 2.5 cm moistsoil. The second half of the nematodes were applied in 10 ml of steriledistilled water on top of the soil surface. Evaluation of insectmortality was performed as previously mentioned.

The experimental design was a randomized complete block with tentreatments and five replications. An analysis of variance and testing ofmain effects and interactions was performed on the insect mortalitydata. Data with zero means were not included in the statisticalanalysis. The data on percentage insect morality (Y) were transformed toarcsin (Y)¹ ²*57.3 and their values expressed in degrees. The originaldata and the transformed data were subjected to an analysis of varianceusing the General Linear Model (GLM) procedure software of SAS (SASInstitute 1991). The protected least significant difference (LSD,P=0.05) procedure was used to compare means of doses and methods ofnematode application onto the soil to control CEW prepupae ingreenhouse.

Inoculum levels and Method of Application of Steinernema. The efficacyof Steinernema riobravis on controlling H. zea prepupae and pupae wasinfluenced by the dose and method used in soil application (Table 3).Since there was a good correspondence in the results of the analysis ofvariance between the original data and transformed data, the results ofthe untransformed data are presented. Data contained in these tableswere rounded after all the calculations have been performed. Theanalysis of variance indicated that there were significant differencesfor both factors dose and method, but their interaction wasnonsignificant (P=0.05). The five doses and two treatment methods werecompared using a mean separation procedure as presented in the two-waytable of Table 3. Table 3 makes the factorial treatment design exhibitand allows all possible pairwise comparisons of these treatments. Themain effect dose and method means are compared because there was nosignificant interaction.

Steinernema riobravis—insect infectivity was a density dependentresponse. The highest rates of infection were obtained with doses of10,000 and 5,000 nematodes/pot (555,555 and 277,778 IJ/m²) whichresulted in 86 and 68% insect infectivity, respectively. The biocontrolobtained with the highest dose was significantly higher than thoseobtained at lower inoculum levels, except with rate of 5,000nematodes/pot (P=0.01) (Table 3). The dependent response. The highestrates of infection were obtained application method also influenced theefficacy of Steinernema riobravis in controlling H. zea prepupae andpupae. The subsurface method resulted in 81% mortality which was higherthan 45% insect mortality achieved with nematodes applied with thesurface method. When nematodes were applied to the soil by thesubsurface method at levels of 5,000 and 10,000 IJ/pot, 90 and 98%insect mortality occurred, respectively (Table 3).

EXAMPLE 4

Field Experiments. The field trials were designed to determine theeffects of dose and timing of nematode application to the soil on thecontrol of H. zea prepupae and pupae. Five concentrations of Steinernemariobravis from Example 1 (0, 25000, 50000, 100000, and 200000nematodes/m²) were applied to the soil at three different time schedulesrelative to the maturity of H. zea larvae infesting the corn ears: 1)when 10% of the H. zea had exited the corn ear to pupate, 2) when 50% ofthe H. zea had developed to large larvae (≧21 mm), and 3) when 40% ofthe H. zea had reached medium larvae (20 mm). Corn plantings betweenfields were separated two weeks from each other. Infective juvenilenematodes stored at 10° C. for 2 wk were suspended in 8 liters of waterand applied with a sprinkling can to each plot at evening time (after1630 hours CST). Control treatments of water were also applied. Afternematode application, prepupae and pupae were extracted with a gardentrowel by carefully scraping the top 10 cm of a 2-m² soil sample in eachplot, at six days after 95% of the H. zea had left the corn ear topupate. Each prepupa or pupa was placed individually in a 20 ml plasticcup, kept in a styrofoam box and processed in the laboratory in the sameday. Evaluation of insect mortality was based on the presence ofSteinernema riobravis nematodes from infected H. zea, as mentioned inthe greenhouse trial of Example 3. The experimental design was arandomized complete block with fifteen treatments and eightreplications. The treatments were arranged in a 5×3 factorial, with fivelevels of nematodes and three times of application already mentioned.Plots were single rows 4m long by 1m wide. Each plot was separated 16mwithin rows and 2m between rows. An analysis of variance and testing ofthe main effects and interactions was performed on the insect mortalitydata. Data transformation, its analysis and mean separation wereperformed using the procedures described in the greenhouse trial ofExample 3.

Inoculum Levels and Timing of Steinernema. The efficacy of Steinernemariobravis was greatly influenced by dose and timing on the control of H.zea prepupae and pupae in corn fields (Table 4). The results ofuntransformed data are presented in Table 4 for reasons previouslymentioned. Analysis of variance indicated significant differences fordose and time but no significant differences for their interactions(P=0.05). Therefore, the main effect means are compared. Generally, eachof these four inoculum levels was significantly greater than the 11%mortality in control plots. Prepupal and pupal infectons were 47, 51,66, and 72% at application rates of 25000, 5000, 100000 and 200000 IJ/m²(Table 4). Natural prepupal and pupal infection in the control plots was11%. The best inocolum level of 200,000 IJ/m² resulted in 72% insectmortality which was significantly (P=0.05) higher than those attained atlower inoculum levels, except with level of 100,000 IJ/m². The besttiming of nematode application occurred when 10% of the H. zea hadexited the corn ear to pupate or when 50% of the H. zea had developed tolarge larvae. This resulted in 77 and 78% insect mortality,respectively. However, poor control of H. zea (22%) resulted from plotsthat received nematodes when 40% of CEW had reached to medium larvae.The highest mortality was obtained in treatments receiving 200,000 IJ/m²applied when 10% of the H. zea had left the corn ear to pupate or when50% of the corn earworms had developed to large larvae. This resulted in95 and 100% of CEW prepupae and pupae infected with Steinernema. Anothersuccessful treatment was with dose of 100,000 IJ/m² applied when 50% ofthe H. zea had developed to large larvae which resulted in 92%infectivity (Table 4).

Residual Efficacy of Steinernema riobravis. A laboratory bioassay andtwo extraction methods were used to determine the residual efficacy andnematode population in soil. The residual efficacy of Steinernemariobravis on the control of H. zea prepupae was performed using a labbioassay method (unpublished data). Soil samples were collected fromcorn plots treated with 200,000 nematodes/m² as in Example 4 on threedifferent dates. Two soil subsamples were collected with a garden trowelfrom the 10-15 cm soil surface of the remaining 2 m² undisturbed soil oneach eight plots each three corn fields. Soil samples were placed inplastic bags, stored in styrofoam boxes and processed in the lab thesame day. Composite soil samples were obtained separately from each ofthe corn fields by gently mixing the soil subsamples. Two aliquants (100cc soil) were transferred each separately into two assay chambers todetect the presence of Steinernema riobravis. Two H. zea prepupae wereburied in soil contained in each ten chambers per field. Chambers wereplaced in dark room (25±2° C.) for five days, then the H. zea wereremoved, rinsed and transferred to “White” trap dishes (White, ibid) todetect the presence of infective juveniles at 12 d after H. zea wereexposed to soil. The residual efficacy of this nematode was estimated bythe percentage of Steinernema—dead insects based on the development ofinfective junveniles in the cadavers of H. zea. Infective juvenilesstarted exiting from infected hosts about 10 d after insects wereexposed to natural soil. Nematodes were collected for theiridentification based on morphological features of IJ and males (Poinar,ibid). The residual population of Steinernema riobravis in soil wasestimated by using the Baermann funnel and the centrifugal flotation(modified) methods described by Barker [Nematode extraction andbioassays. Pp. 19-35. In K. R. Barker, C. C. Carter, and J. N. Sasser,eds. An advanced treatise on Meloidogyne, vol. 2. Methodology. Raleigh:North Carolina State University Graphics, 1985]. Two separate aliquants(100 cc soil) from the composite soil sample of each three corn fieldswere separately processed by the two extraction methods. Nematodes werecollected at 24 and 48 h for the Baermann method. Extracts containingnematodes were counted under a stereoscopic microscopic and a 0.5-mlnematode suspension was added to filter paper (Whatman No. 1) containedin petri dishes for assay of infectivity. One H. zea prepupae was addedto each petri dish (60×15 mm) and incubated in dark room (25±2° C.) for5 d. Then, each dead insect was transferred to White trap dishes (White,ibid) to verify nematode emergence and its identification. Evaluation ofnematode assays were based on numbers of live nematodes per 100 cc soil.In addition to this absolute measure of density, the residual density(Rd) was calculated as a percentage: Rd=(Rp/Tp)*100 where Rp=numbers ofnematodes of a species in a sample; Tp=total number of nematodes of thesame species initially applied to the soil.

Of the two extraction methods, the highest recovery of nematodes fromsoil was obtained with the Baermann funnel (Table 3). Residual densitiesof S. riobravis extracted with the Baermann funnel were 22, 45 and 28%of the total numbers of nematodes applied to the soil on the threedates, respectively. The residual efficacy of S. riobravis as indicatedby the laboratory bioassay resulted in 80, 85 and 90% insect moralityfrom plots that received nematodes 11, 10 and 8 wk after treatment,respectively (Table 5).

EXAMPLE 5

Geographical Distribution. Our research was conducted over 5 consecutiveyears in an irrigated region that extends about 60 km north from the RioGrande River into the U.S.A. and about 60 km south into Mexico. East towest, the region extends from the cities of Brownsville, Tex. andMatamoros, Tamaulipas, Mexico about 190 km to Camargo Tamaulipas. Thisarea is located in the semi-arid subtropics and receives approximately600-700 mm of annual rainfall. Many soil types exist in the area,generally consisting of 25-70% clay, 15-65% sand, 15% silt and a pH<8.About 200,000 ha of irrigated corn are planted annually in this regionin February and early March. Fruiting normally begins in early tomid-May and mature corn earworm and fall armyworm larvae exit the cornto pupate in late May and early June. Little or no pesticide is used tocontrol the corn earworm and fall armyworm larvae that infest this crop.

Sampling Procedure. Quantitative estimates of the number of corn earwormand fall armyworm prepupae and pupae parasitized by the indigenousSteinernema riobravis in fruiting corn was accomplished by excavatingtwo, 1 m² soil samples in each of 90-120 fields annually from 1986 to1990. Samples were taken at 50 and 100 m from field edges after most ofthe larvae had exited the ear to pupate.

Because the standard row spacing for most corn produced in the region is100 cm, the surface area for soil sampled extended from the plants ofone row to the plants of an adjacent row by 100 cm along the rows. Eachsample site was thoroughly searched for corn earworm and fall armywormprepupae, pupae, and pupal exuviae. Insects were initially exposed bycarefully scraping the soil surface with a garden trowel to uncovertunnels leading to the pupae. Following the removal of insects fromexposed pupation chambers, soil was then excavated to a depth of 10-15cm to recover any insects missed in the initial search. All extractedinsects were placed individually in 20 ml plastic cups and transportedto the laboratory for subsequent determination of species, sex, anddevelopmental stage. All dead pupae were examined with the aid of adissecting microscope to determine the presence of adult or infectivejuvenile S. riobravis. Mortality was attributed to the nematode only ifnematodes were observed in the cadaver. We did not attempt to determineother factors resulting in prepual or pupal mortality.

Depending on the year, sampled fields were located on 5 or 6 transectsthrough the main corn-growing area centered near Rio Bravo, Tamaulipas,Mexico and through the center of the irrigation district on the Texasside of the Rio Grande River. The following transects were common to allyears of our study: 1. (west) extended south approximately 25 km fromRio Bravo; 2. (east) located 15 km east of Rio Bravo and extendedapproximately 25 km south; 3. (south) located 15 km south of Rio Bravoand extended from the west side of the irrigation district eastapproximately 25 km; 4. (center) located 5 km south of Rio Bravo andextended from the west side of the irrigation district eastapproximately 25 km; 5. (north) extended from 20 km east to McAllen,Tex. to the east approximately 50 km to Harlingen, Tex. Fields weresampled on an additional transect which extended west from Reynosa toCamargo (ca. 45 km) for two years of the study. From 15 to 20 fields(each separated by ca. 1.5 km) were sampled annually on each transect.

Data analysis. To determine differences in parasitism between species,life stage and year, data were analyzed by analysis of variance andmeans separation was accomplished by computing Least Square Means andtesting the hypothesis, H_(o): LSM (_(i))=LSM(_(j)). Arcsinetransformations were performed on all percentage data before analysis.

Averaged over years, significantly more of the sampled fields harboredcorn earworm prepupae, pupae or exuviae than fall armyworm (92.6 and60.2% respectively; df=1,8 F=7.5 P>F=0.253) (Table 1). Considering onlythose fields where corn earworm or fall armyworm were found in oursamples, 34.2 and 24.2% respectively contained either prepupae or pupaethat were parasitized by Steinernema riobravis. The highest incidence offields with parasitized corn earworm or fall armyworm was observed inyear 1 (52.5 and 36.4% respectively) while the lowest incidence occurredin year 3 (14.1 and 0% respectively).

The highest incidence of corn earworm and fall armyworm parasitism,totaled across prepupae and pupae (including pupal exuviae), occurred inyear 2 (21.3 and 21.2% respectively) while the lowest incidence occurredin year 3 (2.9 and 0% respectively) (Table 7). Although fewer cornearworm prepupae were excavated than pupae and exuviae combined, asignificantly higher percentage of the prepupae were parasitized(df=1,645; F=21.47; P>0.0001). Similarly, a significantly higherpercentage of fall armyworm prepupae were parasitized compared withpupae (df=1,380; F=17.15; P>0.0001). Averaged over all years, 11.6% and9.3% of all corn earworm and fall armyworm excavated were parasitized bySteinernema riobravis respectively. No significant difference in theparasitism rate between corn earworm and fall armyworm was noted.

Corn earworm and fall armyworm parasitism averaged 27.7% and 29.5%respectively, when calculated using only those corn earworm collectedfrom fields where at least one prepupa or pupa was parasitized (Table8). Calculated on this basis, the highest rates of parasitism for cornearworm and fall armyworm (45.3% and 84.6% respectively), were observedin year 2. Averaged over years, there was again no significantdifference win the percentage of parasitism between corn earworm andfall armyworm.

Significant differences in corn earworm parasitism occurred amongtransects (Table 9), when comparing those transects that were common toall years of the study. This comparison included all corn earworm orfall armyworm excavated from fields within transects regardless of thepresences of Steinernema riobravis within individual fields. The highestincidence of corn earworm parasitism (15.4%) was measured in thewestermost transect and the lowest rate of parasitism was observed inthe northernmost transect which was located in the U.S.A. There as nosignificant difference in the rate of parasitism of fall armyworm due totransect location (Table 9).

Considering all prepupae, pupae and pupal exuviae excavated, 23.5% and20.1% of corn earworm and fall armyworm were dead at the time ofcollection (Table 10). The highest mortality occurred in year 2 and thelowest in year 5. When corn earworm mortality was partitioned betweenthe prepupal and pupal stages, a significantly higher percentageoccurred in the pupal stage (80.6%) (df=1,560; F=372, P>0.0001).Similarly, when fall armyworm mortality was partitioned between prepupaeand pupae, a significantly higher percentage occurred in the pupal stage(77.1%) (df=1,262; F=421; P>0.0001). Of those prepupae and pupae thatwere dead at time of collection, 49.4% and 46.1% of corn earworm andfall armyworm, respectively, contained Steinernema riobravis nematodes.The highest percentage of corn earworm and fall armyworm mortalityresulting from parasitism (67.5% and 66.1%, respectively) occurred inyear 1. Averaged over all years, 49.4% and 46.1% of all corn earworm andfall armyworm mortality, respectively, resulted from Steinernemariobravis parasitism. There was no significant difference (P<0.05), inthe percentage of nematode induced mortality between the prepupal andpupal stages.

It is understood that the foregoing detailed description is given merelyby way of illustration anmd that modification and variations may be madetherein without departing from the spirit and scope of the invention.

TABLE 1 Effect of different concentration of Steinernema riobravis onthe mortality of Helicoverpa zea in vitro Insect Probit^(b) NematodesDead mortality LD 95% Fiducial added/prepupa insects^(a) % Value Limits 1  1  5  2 1-3  5  4 20  5 2-7 10  8 40 10  7-13 — — 50 13  9-18 20 1155 15 11-21 40 17 85 48 33-84 80 18 90 65  43-125 — — 95 103   63-231100  20 100  — — ^(a)Based on 20 prepupae. ^(b)The LD₅₀ and LD₉₅ valueswere computed by Probit analysis and added to this table.

TABLE 2 Numbers of Steinernema riobravis nematodes extracted frominfected prepupae and pupae of Helicoverpa zea Average nematodesproduced^(b) Nematodes No. dead Insects^(a) (× 1000) added/prepupaPrepupae Pupae Per prepupa Per pupa  1 0 1  5 2 2 314 (168-460) 190(176-204) 10 1 7 336 (336-336) 317 (200-508) 20 7 4 303 (276-352) 296(292-308) 40 11  6 375 (288-552) 341 (280-452) 80 6 12  257 (200-400)304 (296-384) 100  12  8 326 (112-440) 330 (200-436) ^(a)Based on 20prepupae. ^(b)Range indicated in parenthesis.

TABLE 3 Effects of dose and method of application of Steinernemariobravis onto the soil on the control of H. zea prepupae and pupae ingreenhouse. Nematode Insect Mortality (%) application Dose (No.nematodes/pot) method 0 1250 2500 5000 10000 Mean ^(a) Subsurface 0 5086 90 98 81 a Soil surface 0 28 32 46 74 45 b Mean ^(a) : 0 39c 59b 68ab86a ^(a) Data with zero mean were not included in the analysis. Meansfollowed by a common letter are not significantly different. LSD(Dose) =19.6; LSD(Method) = 13.9; MSE = 466.3; df = 31; (P = 0.05).

TABLE 4 Effects of timing and dose of Steinernema riobravis applied tothe soil on the control of H. zea prepupae and pupae in corn field.Nematode Insect Mortality (%) application Dose: No. nematodes (×1000)/m² time ^(a) 0 25 50 100 200 Mean ^(bc) 10% cutouts 20 68 68 76 95 77a 50% large larvae 14 43 77 92 100 78a 40% medium larvae  0 29  829  21 22b Mean ^(c) : 11c 47b 51b 66ab  72a ^(a) Timing nematodeapplication when 10% of CEW had exited the corn ear to pupate (cutouts)and when the corn fields were infested with large or medium larvae. ^(b)Data for dose zero were not included in the Time means. ^(c) Time anddose means followed by a common letter are not significantly different.LSD(Dose) = 18; LSD(Time) = 14; (P = 0.05).

TABLE 5 Residual efficacy of Steinernema riobravis against H. zeaprepupae after nematode application to corn field plots as determned bytwo extraction methods and a laboratory bioassay. Nematode Number ofnematodes/100 cc soil Insect application Baermann Rd ^(a) Centrifugal Rdmortality (%) period funnel (%) flotation (%) Lab bioassay ^(b) 1 43 2210  5 80 2 90 45 39 20 85 3 56 28 26 13 90 Mean: 63 32 25 13 85 ^(a) Rd,residual density of nematodes is the ratio of the number of nematodesextracted to total number of nematodes initially applied to the soil(200 IJ/100 cc soil). ^(b) Based on 20 H. zea prepupae.

TABLE 6 Percentage of corn fields in Lower Rio Grande Valley whereexcavated corn earworm and fall armyworm were parasitized by Steinernemariobravis. Number Percent fields Number fields with with parasitized^(a)Year fields sampled cew faw cew faw^(b) 1 120 118  99 52.5 36.4 2 105 8337 37.4 21.6 3  90 85 17 14.1  0.0 4 100 99 87 28.3 26.6 5 100 92 7030.4 11.4 Total 515 477  310  Average 34.2 24.2 ^(a)Based on fieldswhere pupae were collected from soil samples. ^(b)cew = H. zea; faw = S.frugiperda

TABLE 7 Incidence of H. zea and S. frugiperda prepupae and pupaeparasitism by Steinernema riobravis in corn fields in the Lower RioGrande Valley. Percent parasitized^(a) prepupae pupae total^(b) Year cewfaw cew faw cew faw^(c) 1 49.1 (57) 74.4 (39) 17.7 (813) 15.7 (549)19.8a 19.6a 2 25.0 (20) 60.0 (5) 21.0 (205) 19.1 (47) 21.3a 21.2a 3  4.5(88)  0.0 (5)  2.6 (431)  0.0 (26)  2.9bc  0.0c 4 11.0 (82)  2.3 (177) 7.0 (740)  3.6 (754)  7.4bc  3.3b 5 18.0 (61) 14.3 (7)  8.3 (539)  4.7(193)  9.3b  5.0b Avg. 18.5 15.5 10.8  8.3 11.6  9.3 ^(a)Numbers inparenthesis indicate number of insects observed. ^(b)Column valuesfollowed by the same letter are not significantly different (p<.05) asdetermined by Least-squares means. ^(c)cew = H. zea; faw = S. frugiperda

TABLE 8 Frequency of H. zea and S. frugiperda parasitism by Steinernemariobravis in fruiting corn fields where at least one parasitizedprepupae or pupae was excavated. Number Number insects Percent fieldsexcavated parasitized^(a) Year cew faw cew faw cew faw 1  62 36  549 31632.1b 37.6bc 2  31  8  115  13 45.3a 84.6a 3  12  0  87  0 17.9bc  — 4 28 11  334 407 19.6c 13.7c 5  28  8  243  21 23.8bc 47.6b Total 161 631328 757 Average 27.7 29.5 ^(a)Column values followed by the same letterare not significantly different (p<.05) as determined by Least-squaresmeans. ^(b)cew = earworm; faw = fall armyworm

TABLE 9 Spatial distribution of corn earworm and fall armywormparasitized by Steinernema riobravis in the Lower Rio Grande Valley^(a).Number of Percent fields parasitism^(b) Transect cew faw cew faw^(c)West 85 54 15.4 a  9.4 a South 89 58 12.8 a  6.8 a East 83 53 10.1 ab18.2 a Center 84 60  9.8 b  3.3 a North 81 48  8.6 b  6.3 a ^(a)Datafrom fields along predetermined transects. ^(b)Column values followed bythe same letter are not significantly different (p<.05) as determined byLeast-squares means. ^(c)cew = H. zea; faw = S. frugiperda

TABLE 10 Percentage mortality of H. zea and S. frugiperda excavated fromfruiting corn fields in the Lower Rio Grande Valley resulting fromparasitism by Steinernema riobravis. Percent of mortality Dead resultingfrom number percent Steinernema riobravis ^(a) Year cew faw cew faw cewfaw^(b) 1 255 174 29.3 29.6 67.5 a 66.1 a 2  74  23 32.9 44.2 64.9 ab47.8 ab 3 111  6 21.4 19.4 13.5 c  0.0 c 4 176 131 21.4 14.1 34.7 c 23.7c 5  96  28 16.0 14.0 58.3 b 35.7 b Total 712 362 Average 23.5 20.1 49.446.1 ^(a)Column values followed by the same letter are not significantlydifferent (P<.05) as determined by Leasts-squares means. ^(b)cew = H.zea; faw = S. frugiperda

We claim:
 1. A composition for use as a biopesticide comprising aninsecticidally effective amount of entomopathogenic Steinernemariobravis American Type Culture Collection accession no. 75727 which isisolated from the environment and an inert carrier.
 2. The compositionas described in claim 1 wherein said carrier comprises water in whichsaid Steinernema riobravis are suspended.
 3. The composition asdescribed in claim 1 wherein said carrier is a solid phase material andsaid Steinernema riobravis are immobilized on or within said carrier. 4.The composition as described in claim 3 wherein said carrier comprises ahydrogel agent.
 5. The composition as described in claim 4 wherein saidhydrogel agent comprises alginate.
 6. The composition as described inclaim 3 further comprising a humectant.
 7. The composition as describedin claim 7 wherein said carrier comprises an encapsulating agent andsaid Steinernema riobravis are encapsulated within said agent.
 8. Thecomposition as described in claim 1 wherein said Steinernema riobravisare produced by in vitro cultivation.
 9. The composition as described inclaim 1 wherein said Steinernema riobravis are produced by in vivocultivation in a host insect and recovered therefrom.
 10. Thecomposition as described in claim 1 wherein said Steinernema riobravisare isolated and substantially pure.
 11. The composition as described inclaim 1 wherein said Steinernema riobravis are isolated in pure form.